Ribonuclease (RNase) enzymes degrade polymeric ribonucleic acids (RNA) into shorter fragments or component nucleotides. All organisms produce ribonucleases and these enzymes are found in most environments. The properties of a number of ribonucleases are described by D'Alessio and Riordan (1997. As a group, most ribonucleases are specific for single-stranded RNA and will not cleave RNA in duplex form. Further, ribonucleases generally cleave at the 3′-end of a ribonucleic acid phosphodiester linkage. Many different RNase enzymes exist, some of which have little or no substrate preference while others are sequence specific. For example, ribonuclease I, from E. coli, is a non-specific endoribonuclease that degrades RNA by cleavage at any base. Ribonuclease A, from mammalian pancreas, is a base-specific endoribonuclease that degrades RNA by cleavage following a pyrimidine (uridine or cytosine) base. Ribonuclease T1, from Aspergillus oryzae, is a base-specific endoribonuclease that degrades RNA by cleavage following a guanosine residue. These three RNase enzymes are noteworthy in that they are routinely employed in standard molecular biology protocols to remove unwanted RNA from samples or as a component in certain assay procedures.
Single-strand specific RNases are the primary nuclease activity encountered in research laboratories as an unwanted contaminant. Double-strand specific RNases have been described, however these are rare and not routinely found in most laboratory settings. RNase H cleaves RNA only when complexed as a heteroduplex with DNA and is not of concern as a laboratory contaminant.
Ribonucleases are present in all laboratories as ubiquitous environmental contaminants. RNases are also found in most molecular biology laboratories as purified enzyme stocks. In laboratories that study RNA, careful attention to experimental protocol is needed to avoid contamination of reagents with RNases; for example, gloves must be worn at all times to prevent contact with the RNases that are universally present on human skin. Regardless of source, the presence of a contaminant RNase will degrade any RNA that comes in contact with that reagent, resulting in the loss of valuable samples or interfering with time-consuming experiments. Once present, removing RNase activity from a laboratory reagent is difficult. Most RNase enzymes are remarkably stable and survive harsh treatments that are routinely used to eliminate other unwanted biologic activities, such as autoclaving. Methods that remove RNase activity range from baking glassware at very high temperature to treating reagent stocks with the highly toxic chemical diethylpyrocarbonate (Sambrook et al., 1989). In spite of such attention, RNase contamination remains a chronic problem and monitoring for the presence of RNase activity is a routine quality control (QC) step in most research and industrial laboratories. As such, methods are needed that would detect the presence of RNase activities commonly encountered in the laboratory setting and that are suitable for routine, frequent use.
Many methods have been devised attempting to measure RNase activity. RNase assays can be grossly divided into methods that detect degradation of heterogeneous RNA obtained from biological sources and methods that detect specific cleavage of a well-defined synthetic substrate, such as an oligonucleotide. In general, use of a synthetic substrate affords both increased sensitivity and improved specificity. Many different detection modalities have been incorporated into these assays, including direct staining, spectrophotometric and colorimetric readouts, chromogenic cascade, radioactive tracer, fluorescence polarization, and fluorescence quenching methods.
Choice of detection method will affect assay sensitivity and ease of use. For use in determining the presence or absence of RNase contamination in laboratory reagents, the method should be sufficiently sensitive to detect the presence of RNase enzymes at the lowest level that will degrade experimental samples in actual use. An insensitive assay would “pass” reagents that are contaminated, which is undesirable. Conversely, an assay could be too sensitive and might “fail” reagents that, from a practical standpoint, are not contaminated and would therefore also be undesirable.
A detection limit within the range of 1-100 picogram/ml (pg/ml) of RNase A is ideal for a reagent QC assay. Commercial assays currently available are sensitive in the 10-100 pg/ml range (Ambion Catalog, 1999). Since such an assay would be used repeatedly, it is also desirable that the method be rapid and easy to perform. Preferably, such an assay could be done at the site of suspected contamination and offer a rapid visual readout.
The original unit definition of ribonuclease activity is based upon the method of Kunitz (1946) which employs a spectrophotometric assay to measure the decrease in absorbance at 300 nm that occurs with degradation of heterogeneous RNA. While the method has been improved (Oshima, 1976), it is insensitive and therefore of little use as a quality control (QC) assay.
Another method to detect RNase activity involves separation and assay of component enzyme activities within a sample using polyacrylamide gel electrophoresis (Wilson, 1969). RNase enzymes can be detected in the acrylamide matrix by direct staining or by incubation with a heterogeneous substrate RNA and an RNA staining dye, such as toluidine blue. While conceptually simple, this approach is time-consuming and relatively insensitive, having a lower limit of detection of about 1 unit of RNase I. In an improvement of this technique, Karpetsky (1980) describes a polynucleotide/polyacrylamide-gel electrophoresis method that improves sensitivity to below 100 pg of RNase A. However, even the improved method remains slow and cumbersome and is better suited to the analysis of ribonuclease activities in biologic specimens than to the QC of laboratory reagents.
Another approach to detect RNase activity is described by Egly and Kempf (1976). This procedure detects release of soluble 125Iodine-labeled RNA from an insoluble RNA-agarose matrix in the presence of ribonuclease. The method is capable of detecting the presence of RNase A at levels as low as 0.01 pg/ml and is actually too sensitive for use as a routine QC assay. Furthermore, this method employs a hazardous radioactive isotope as reporter that is not desirable for use in most laboratory or industrial settings.
Another approach to detect RNase activity is described by Wagner (1983). RNA forms a complex with Pyronine-Y that has an optical absorbance maximum at 572 nm. Degradation of high molecular weight RNA by ribonuclease activity results in loss of absorbance at 572 nm in a linear and quantitative fashion. The method, however, is only capable of detecting about 2 ng/ml RNase A in a test sample and has insufficient sensitivity for use as a QC assay.
Another approach to detect RNase activity was described by Greiner-Stoeffele (1996). The dye methylene blue intercalates into high molecular weight ribonucleic acid forming a dye-RNA complex. Upon degradation by ribonuclease action, methylene blue is released and absorbance at 688 nm decreases. This method, however, is also relatively insensitive and can detect ribonuclease activity only down to about 25 ng/ml, which is inadequate for use as a QC assay.
Another approach to detect RNase activity is described by Karn (1979). Ribonuclease A-mediated cleavage of a synthetic ribonucleotide dimer substrate was detected by a cascade of enzymatic reactions involving adenosine deaminase, nucleoside phosphorylase, and xanthine oxidase that ultimately forms a detectable blue tetrazolium salt. The method can detect the presence of 0.066 units of RNase A (about 100 ng), insufficient for use as a QC assay. Furthermore, the procedure is lengthy, complex and requires modification to detect the presence of ribonucleases other than RNase A.
Another approach to detect RNase activity is described by Witmer (1991). A synthetic ribonucleotide substrate, U-3′-BCIP, was synthesized that releases a reporter group in the presence of RNase A that could be detected spectrophotometrically by absorbance at 650 nm. While this chromogenic method is simple to use, it is insensitive and is better suited for applications such as the in vivo bacterial colony assays taught by Witmer than for use as a reagent QC assay.
Fluorescence-quenching detection is used in many applications in the biological sciences; representative examples include methods to detect proteolytic enzyme activity (Yaron, 1979), methods to detect DNA restriction endonuclease activity (Ghosh, 1994), methods to detect the 5′-nuclease activity of DNA polymerase (Gelfand, 1993), methods to detect nucleic acid sequence identity (Gelfand, 1993; Tyagi, 1999; Livak, 1999; Nazarenko, 1999; Nadeau, 1999), and methods to detect bimolecular protein interactions in an immunoassay (Maggio, 1980). A synthetic oligoribonucleotide having a Fam reporter group and a Tamra quencher group has been used as a FRET probe to detect hammerhead ribozyme activity (Hanne, 1998). Fluorescence Resonance Energy Transfer (FRET) and fluorescence quenching methods are reviewed by Morrison (1992).
Zelenko (1994) describes synthesis of a dinucleotide substrate uridylyl-3′5′-deoxyadenosine that is conjugated to a fluorophore (O-aminobenzoic acid) on one end and a fluorescence quencher (2,4-nitroaniline) on the opposite end of the molecule. Cleavage by RNase A separates the fluorophore and quencher, leading to a detectable increase in fluorescence. The substrate was designed specifically for use in kinetic studies of RNase A activity and will react only with the subset of ribonuclease enzymes that cleave at a uracil residue. Having a limited spectrum of sensitivity, this reagent is not suitable for use as a single substrate in an RNase QC assay.
James (1998) describe an alternative substrate for kinetic studies of RNase A in which a 9-mer chimeric oligonucleotide that contains a single ribonucleotide uracil base flanked by deoxyadenosine residues was modified with a 5′ fluorescein reporter group and a 3′ rhodamine quencher group. The utility of the substrate is limited in that it can detect only those ribonucleases that cleave at a uracil residue. Further, assay results must be detected using a fluorometer due to background fluorescence of the rhodamine quencher group.
Kelemen (1999) describes a similar substrate having somewhat greater sensitivity measuring RNase A kinetics with the following composition: SEQ ID. NO:2: 5′ Fluorescein-AuAA-Tamra 3′. Like the James reagent, the Kelemen substrate is limited to detecting ribonucleases that cleave at a uracil residue and requires the use of a fluorometer.
James (1998) and Kelemen (1999), therefore, have described use of fluorescent-labeled oligonucleotide probes with FRET/quenching to study the catalytic properties of RNase A. Both compositions are chimeric DNA-RNA oligonucleotides that contain a single internal uridine base, use a fluorescein dye as reporter group, and use a quencher group that is a fluorophore that itself emits light in the visible spectrum, so methods that use these substrates require availability of a fluorometer for detection. These probes were optimized for kinetic studies of RNase A and cannot be used to detect the presence of RNase enzymes that do not cleave at a uridine residue. In addition, both compositions include DNA residues, which are subject to cleavage by DNase enzymes, so cleavage is not RNase specific. They are, therefore, not useful as a tool to assay for the presence of contaminating RNase activity.
Burke (1998) describes a method that utilizes fluorescence polarization detection techniques to measure cleavage of short, synthetic nucleic acid probes. A commercial kit for performing RNase detection of Burke is available (Pan Vera Catalog, 2000). Wilson (2000) describes a variant of this technique that examines real-time degradation of a long, synthetic RNA species (made by in vitro transcription) using fluorescence anisotropy. The fluorescence polarization-based techniques that must be employed, however, cannot be performed without a specialized fluorescence polarization fluorometer, which is not available in most laboratories.
Another commercial kit for the detection of RNase activity measures the release of soluble fluorescent dye from a precipitated (i.e., insoluble) fluorescent RNA substrate (Pan Vera Catalog, 2000). This method is less sensitive than the fluorescence polarization method and also requires availability of a fluorometer, thereby limiting the utility of the assay.
A commercial kit is available that uses a biotin-labeled RNA substrate immobilized on dipsticks to test for the presence of RNase activity (Ambion Catalog, 1999). Detection is achieved using a visual colorimetric method. In the absence of RNase, the substrate remains intact and the calorimetric assay develops a blue spot on the dipstick while in the presence of RNase the label is cleaved and no color develops. This assay is labor intensive, takes over 3 hours to perform, and is not well suited for high-throughput QC use.
Another commercial RNase detection kit employs gel electrophoresis to visualize degradation of a high molecular weight RNA in the presence of ribonuclease activity (Mo Bio, Web Catalog, 2000). The method is a multi-step, labor intensive protocol that is very expensive, making it unsuitable for routine QC use.
It is apparent from the above discussion that, while progress has been made in methodology to detect ribonuclease activity, existing assays have significant limitations. None are suitable for use as a universal ribonuclease detection system (i.e., a QC assay). A ribonuclease detection method suitable for use in a QC assay should meet the following 7 criteria:                1) The assay will be highly sensitive.        2) The assay will be highly specific.        3) The assay will detect a broad spectrum of ribonuclease activities.        4) The assay reagent(s) will be inexpensive and suitable for commercial manufacture.        5) The assay method will be both simple and rapid.        6) The assay method will allow for visual detection and will not require the use of highly specialized equipment.        7) The assay will not employ any hazardous compounds.        
Clearly new methods, or improvements in earlier methods, are needed. In particular, a need exists for an RNase assay that is rapid, sensitive, within the desired range and allows for visual detection.